production of biodiesel-like components by the type i
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Clemson UniversityTigerPrints
All Theses Theses
8-2013
Production of biodiesel-like components by theType I methanotroph Methylomonas methanicaMegan Diane BurdetteClemson University
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Recommended CitationBurdette, Megan Diane, "Production of biodiesel-like components by the Type I methanotroph Methylomonas methanica" (2013). AllTheses. 2309.https://tigerprints.clemson.edu/all_theses/2309
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Production of biodiesel-like components by the Type I methanotroph Methylomonas methanica
A Thesis Presented to
the Graduate School of Clemson University
In Partial Fulfillment of the Requirements for the Degree
Master of Science Microbiology
by Megan Dianne Burdette
August 2013
Accepted by: Dr. J. Michael Henson
Dr. Jeremy Tzeng Dr. Sarah Harcum
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ABSTRACT
Bacteria that utilize methane as a sole source of carbon and energy are referred
to as methanotrophs. Industrial uses of these types of organisms include the
production of poly-hydroxybutyrate as well as the degradation of some
chlorinated hydrocarbons that are considered pollutants in the environment.
Methanotrophs also play an important role in methane cycling in the
environment. Methane gas has the ability to trap 20 times more heat than
carbon dioxide, which makes it a potent greenhouse gas. A defining
characteristic of methanotrophs is the production of extensive intracytoplasmic
membranes composed of lipids that are 16 or 18 carbons in length, similar to
those of soy biodiesel. Therefore, considering the existing abundant supply of
methane gas, another potential industrial application of these organisms is to
utilize the intracytoplasmic membrane lipids as a source of components similar to
biodiesel. In this study, we hypothesized that by optimizing the growth
temperature as well as the copper concentration, a greater amount of desired
lipids would be produced. Results indicated that the production of total lipids and
specifically the fatty acid methyl ester (FAME)16:1 was greater at 25 °C while
lower amounts of 16:1 were produced at 30 °C and 33 °C. Bacterial growth was
not observed at 20 °C and the bacterial cells clumped together at 35 °C. M.
methanica was then grown at 25 °C in the presence of six concentrations of
copper ranging from 0 to 50 µM, with 5 µM yielding the highest production of
16:1. No growth occurred with 50 µM copper at 25 °C. Based on these results,
M. methanica was grown in a 2 L flask at 5 µM copper at 25 °C. The weight of
total biodiesel-like lipids under these growth conditions was 4.8% of the total
biomass, with C16:1 comprising 70% of the total FAMES. Results of this
research indicate that temperature affects the lipid profile and that copper
concentration affects the amount of lipids produced. Thus, it is possible for M.
methanica to produce biodiesel-like lipids from methane gas.
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DEDICATION
I would like to dedicate this work to my parents, Michael and Dianne
Burdette, as well as my sister, Kylie Burdette. Thank you for your love, support,
and encouragement.
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ACKNOWLEDGMENTS
I would like to acknowledge my advisor, Dr. J. Michael Henson for his
guidance. Also, I would like to thank my fellow lab members: Abhiney Jain,
Sandra Bediako, and Ryan Hammonds.
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TABLE OF CONTENTS
Page
TITLE PAGE ........................................................................................................i ABSTRACT.........................................................................................................ii DEDICATION..................................................................................................... iii ACKNOWLEDGMENTS ....................................................................................iv LIST OF TABLES..............................................................................................vii LIST OF FIGURES .......................................................................................... viii CHAPTER I. Introduction ........................................................................................ 1 Ecology of Methanotrophs............................................................ 1 Production of Biodiesel from Microorganisms .............................. 6 Methane Metabolism .................................................................. 12 II. Materials and Methods .................................................................... 17 Materials..................................................................................... 17 Methods...................................................................................... 18 Experimental Design .................................................................. 20 III. Results and Discussion ................................................................... 26 IV. Conclusion ....................................................................................... 53 APPENDICES.................................................................................................. 58 A: Methane and FAME Standards ....................................................... 59 REFERENCES ................................................................................................ 54
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LIST OF TABLES
1 Characteristics of Types I, II and X methanotrophs........................... 3 2 Lipid Yield of M. methanica at late-exponential and stationary phases, temperatures 25°C, 30°C, and 33°C ......................................................................................... 42 3 Lipid yield of M. methanica at different copper concentrations. Culture conditions are 25°C, late-exponential phase..................................................................... 49 4 Lipid yield and carbon balance in 2 liter flask .................................. 51
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LIST OF FIGURES
Figure 1. The quantities of biomass and FAMES produced
during growth of M. methanica at 30°C and the quantity of methane gas consumed during growth..................... 29
Figure 2. Growth Profiles of M. methanica at (a) 25°C
(b) 30°C and (c) 33°C………………………………………………32 Figure 3. Methane consumption during growth at (a)25°C (b) 30°C and (c) 33°C................................................................. 33 Figure 4. Plot used to normalize dried biomass to cell density.................. 35 Figure 5. The quantity of biomass and percentage total FAMEs produced by M. methanica at late-exponential and stationary phases while growing at temperatures 25°C, 30°C, and 33°C. ............................................................... 37 Figure 6. The quantity of total and individual FAMEs produced by M. methanica while growing at exponential and stationary phases, 25°C, 30°C, and 33°C .................................. 40 Figure 7. The ratios of FAME produced by M. methanica
while growing at late exponential and stationary phase and at the temperatures of 25°C, 30°C,
and 33°C. ................................................................................... 41 Figure 8. The amount of total biomass and percentage of total FAMEs produced when M. methanica was grown at
different copperconcentrations…………………………………….46 Figure 9. The quantity of total and individual FAMEs produced by M. methanica while growing at copper concentrations 0 µM, 1 µM, 5 µM, 10 µM, 20 µM ...……………………………….47
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Figure 10. The ratios of FAMEs produced by M. methanica when grown with different copper concentrations at 25°C during late-exponential phase.. .................................... 48 Figure 11. Amount of total lipids and percentage of individual lipids produced in 2 liter flask ..................................................... 50 Figure 12. Ratio of individual lipids in 2-L reactor........................................ 51 Figure 13. The quantities of biomass and FAMES produced during growth of M. methanica at 25°C, late-exponential phase, 5 µM copper and the quantity of methane gas consumed during growth. ........................................................... 52
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CHAPTER ONE
Introduction
1.1 Ecology of Methanotrophs Methanotrophic bacteria are a unique group of gram-negative bacteria that are
characterized by the ability to use methane as a single source of carbon and
energy. Methanotrophs are a subset of bacteria called methylotrophs that are
capable of oxidizing a variety of single carbon compounds as sources of carbon
and energy. Generally, methanotrophs reside in aerobic environments, and are
members of the Proteobacteria phylum.
Methanotrophs occur naturally in soil, sediments of marshes and swamps,
eutrophic lakes, rice paddies, vegetated wetlands, aquifers and ocean waters
(Hanson and Hanson 1996). The first methanotroph was isolated in 1906 by
Söhngen (Hanson and Hanson 1996) who noticed that environments such as
swamps and eutrophic lakes produced a large amount of methane, but a smaller
amount was present in the atmosphere. He speculated that the reason for this
inconsistency was methane-oxidizing bacteria, and he isolated the first
methanotroph, Bacillus methanica, from a freshwater lake (Söhngen 1906). This
bacterium was later renamed Methylomonas methanica, and was categorized
with over 100 new methanotrophic isolates by Whittenbury et al. (1970). Hanson
further enhanced the taxonomic classifications developed by Whittenbury by
examining the ecology of methanotrophs (Hanson 1980).
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Since the extensive research performed by Whittenbury et al. (1970),
methanotrophic bacteria have been isolated from acidic peat bogs, thermophillic
environments such as hot springs, alkaliphylic soda lakes, and psychrophilic
environments in Siberia and Antarctica. The use of molecular biology techniques
made possible the detection of methanotrophs in these extreme environments.
Methanotrophs are divided in to three groups based on cell structures and
metabolic capacity: Type I, Type II, and Type X (Table 1). The metabolic basis
for these three different groups is the different formaldehyde assimilation
pathways as well as differences in intracytoplasmic membrane structure. Type I
methanotrophs include the genera Methylocaldum, Methylosphaera,
Methylomicrobium, Methylomonas and Methylobacter. As members of the γ-
Proteobacteria, these organisms utilize the ribulose monophosphate (RuMP)
pathway to assimilate formaldehye into cellular material. Type I organisms
possess intracytoplasmic membranes that are arranged in bundles, and are
predominately composed of 16:1 fatty acids. Alternatively, Type II
methanotrophs include the two genera of Methylosinus and Methylocystis.
These organisms are members of the α-Proteobacteria family, and use the serine
pathway for formaldehyde assimilation. Type II methanotrophs are characterized
by intracytoplasmic membranes arranged around the periphery of the cell, and
contain fatty acids that are 18 carbons in length. Finally, Type X methanotrophs
like genus Methylococcus, use the RuMP pathway for formaldehyde assimilation
as well as small amounts of ribulose-bisphosphate carboxylase, an enzyme that
3
is present in the serine pathway. These methantrophs also grow at higher
temperatures than Types I and II methanotrophs. A model organism for Type X
methanotrophs is Methanococcus capsulatus (Bowman 2006).
Table 1. Characteristics of Types I, II and X methanotrophs.
aSymbols: +, 90% or more of strains are positive; and −, 90% or more of strains are negative. RuMP pathway, ribulose monophosphate pathway. bAbsent in most Type I methanotrophs but is present in some strains of Methylococcus and Methylomonas. Adapted from (Bowman 2006)
Characteristics Type I Type X Type II
Genus
Methylococcaceae Methylosphaera Methylobacter Methylomicrobium Methylomonas
Methylococcus Methylocaldum
Methylocystaceae Methylosinus Methylocystis
Intracytoplasmic membrane arrangement
Disks Disks Peripheral
Major fatty acid carbon chain length
16 16 18
Soluble methane monooxygenase b
−a - +
Carbon assimilation pathway
RuMP RuMP Serine
Mol% G+C (Tm) 43-60 56-65 60-67 Phylogenetic group (Proteobacteria)
Gamma Gamma Alpha
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Methanotrophic bacteria play an important role in global carbon cycling by
oxidizing methane gas produced by methanogenic archaea before it enters the
atmosphere. For this reason, methanotrophs are practically ubiquitous in surface
sediment samples and thrive in areas where supplies of methane and oxygen are
readily available. In freshwater environments, research has indicated that
methanotrophs comprise a significant proportion of the total bacterial biomass,
and are thought to be responsible for up to 50% of the annual methane flux
(Cicerone and Oremland 1988). A study of methane cycling in Lake Washington
revealed that methane oxidation occurred in the oxygenated portion of the top 6-
7 mm of the lake sediments, and the rate of oxidation was controlled by the
concentration of methane (Kuivila et al. 1988). Because of the integral role they
play in global carbon cycling, methanotrophs represent a basic trophic level of
the food web. Studies utilizing 13C-labeled isotopes have been successful in
tracing carbon derived from methanotrophs to various higher trophic levels of
aquatic life (Boschker et al. 1998). Estuarine and ocean environments have also
been thought to harbor methanotrophic populations. Monitoring of ocean surface
waters has revealed a super saturation of methane when compared to
atmospheric levels, therefore a biological oxidation of methane was suspected
(Lidstrom 2006). This observation has just recently been confirmed using 16S
rRNA gene cloning and restrictive fragment length polymorphism analysis,
indicating the presence of numerous methanotrophic species (Lidstrom, 2006).
The use of these molecular techniques revealed a broad range of
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methanotrophic 16S rRNA gene sequences that represented both Type I and
Type II organisms (McDonald et al. 2005). Other molecular techniques such as
denaturing gradient gel electrophoresis (DGGE) and quantitative PCR (qPCR) in
conjunction with enzyme specific primers are being used to analyze the
composition of methanotrophic communities as well as identify individual
organisms in environments such as peat bogs, rice paddies, and eutrophic lakes
(Lidstrom 2006).
1.2 Production of biodiesel from microorganisms
The use of plant products to power vehicles was first introduced by Rudolph
Diesel at the 1900 World’s Fair (Hanson and Hanson 1996). Diesel
demonstrated that his invention, the diesel engine, could operate on 100%
peanut oil. After the introduction of this concept, the transesterification of plant-
derived oils was developed in order to produce biodiesel (Chisti 2007). The use
of plant-derived biodiesel was soon replaced by diesel fuel produced from fossil
fuels because of limited availability. However, the need for an alternative source
of fossil-fuel as the source of diesel fuel has recently become crucial in order to
reduce greenhouse gas emissions and dependence on foreign sources of crude
oil.
Traditionally, biodiesel has been produced from plant-derived oils such as
corn and soybean. However, many of these plants are also primary edible
agricultural crops, and this conflict of use places a strain on the agricultural
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industry. This dilemma has driven the search for alternative sources of biodiesel
components, which includes microorganisms.
The current production of biodiesel from microorganisms is based on the
growth of photosynthetic or heterotrophic microalgae. Algae are reported to
produce 60% lipids per cell (wgt/wgt) (Drapcho et al. 2008). The algae used for
this photosynthetic process are autotrophic, thus requiring large amounts of land
for cultivation because of their requirement for sunlight. Thus, the use of
photosynthetic algae sources may not be economically or environmentally
feasible for long-term biodiesel production. The most common growth substrates
used in the heterotrophic process are sugars derived from corn and sugarcane.
Even though the cultivation of in algae in this manner produces high yields of
biomass, the process of delignification and saccharification of cellulose to
glucose is still required for the algae to use these substrates as a source of
energy.
While methanotrophic bacteria produce a lower percentage of membrane
lipids per cell (wgt/wgt), an advantage is present in the fact in that methanotrophs
utilize methane gas, which is also a greenhouse gas, as sole source of carbon
and electrons. Methanotrophs use the enzyme methane monooxygenase as a
catalyst to oxidize methane. Man-made catalysts require large amounts of heat
and expensive materials to function (Balasubramanian et al. 2010). Using MMO
as a catalyst to extract the energy stored in methane molecules may be efficient
and less costly than chemical catalysts.
7
Methane is the second most abundant organic gas present in the
atmosphere. The concentration of atmospheric methane has more than doubled
from 750 parts per billion (ppb) in the 18th century to 1750 ppb present day.
Each year an estimated 810 Tg (1 Tg = 1 x 1012 grams) of methane are produced
by natural and anthropogenic sources. Wetland areas are a major source of
environmental methane, producing 174 Tg of methane per year. These natural
habitats are characterized by standing water and sediment, which provides an
ideal growth environment for anaerobic methanogenic archaea. A significant
amount of methane, 22 Tg per year, is also produced in the hindgut of termites.
Additional sources of naturally produced methane include methane hydrates in
oceans (15 Tg per year) and methane released from geological activities
(9 Tg per year) (Smith 2012). In 2010, approximately 10 percent of all human
related greenhouse gas emissions consisted of methane gas. These
anthropogenic sources include methane produced by anaerobic decomposition
of waste materials in landfills (54 Tg), and rice cultivation (54 Tg). Another human
influenced activity is the production of methane by cows on dairy farms (84 Tg).
Methanogenic archaea reside in the rumen of cattle, and serve the purpose of
assisting in the conversion of grass to more easily digested compounds (Smith
2012). Furthermore, 36 Tg of methane gas is released from coal beds during
coal mining, and the burning of agricultural biomass releases 47 Tg of methane
per year. Finally, 61 Tg of methane is released into the atmosphere as a result
of activities associated with industry (Smith 2012). Methane is also the primary
8
component of natural gas, and 144.7 Tg of methane are released from natural
gas pipelines every year. In addition to these sources, the burning of fossil fuels
and other human related activities produces a significant amount of methane
(Smith 2012).
The US EPA provides information regarding greenhouse gasses such as
carbon dioxide and methane to indicate their global warming potential ratings
(US EPA 2010). This calculated number is the total amount of energy that a
specific gas is capable of absorbing over a period of 100 years. While methane
has a shorter lifetime in the atmosphere than carbon dioxide, its global warming
potential is 21, which is much greater than that of carbon dioxide which has a
global warming potential of 1. Methane has the ability to trap infrared radiation
more efficiently than carbon dioxide, and therefore contributes to global warming
about 20 more times than carbon dioxide (US EPA 2010).
Studies have shown that methane-oxidizing bacteria are crucial in the
global methane cycle because they consume the majority of methane produced
by wetlands and rice paddies before it can be released into the atmosphere
(Hanson and Hanson 1996). Considering the amount of methane produced by
landfills, wastewater treatment plants, and the abundant supply of natural gas,
methane gas should be considered a viable source of energy. The conversion
of a readily available, renewable resource into liquid biodiesel would be beneficial
both to the economy as well as the environment.
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Globally, industry as well as typically everyday life is largely dependent
upon fossil fuels. In 2011, 3.19 billion barrels of crude oil were purchased and
consumed in the United States alone (US EIA 2012). Multiple products can be
derived from each barrel of oil, which is approximately 42 gallons. The
predominant product is gasoline, which is approximately 19 gallons per barrel,
followed by diesel fuel, which is approximately 11 gallons per barrel
(US EIA, 2011). In the United States, gasoline is the most predominant fuel, and
it powers 254 million vehicles. On a larger scale, the United States consumes 44
percent of gasoline worldwide (Horton, 2008). Alternatively, the primary fossil
fuel in Europe is diesel. The popularity of the diesel automobile has increased
because of the tax incentives when purchasing diesel fuel as well as an increase
in fuel mileage and efficiency in new diesel engines (Webster, 2009). Gasoline
prices in Europe are also significantly higher than those in the United States,
costing up to $8.50 per gallon, which is nearly double the average price in the
US.
As stated previously, diesel fuel is derived from crude oil. It has a greater
density and boiling point than gasoline, and it is composed of hydrocarbons
ranging from 11 to 22 carbons in length. The hydrocarbons present in diesel are
either paraffin or aromatic. Paraffin hydrocarbons are straight chains of carbons,
and aromatic hydrocarbons have carbons attached to a benzene ring.
Comparatively, soy biodiesel is composed of lipids extracted from soybeans.
These lipids are converted to fatty acid methyl esters (FAMEs) ranging from 16 to
10
18 carbons in length. Methylomonas methanica, the organism chosen for this
study, produced extensive cytoplasmic membrane lipids that, when converted to
FAMEs, are 14, 16, and 18 carbons in length. Although FAMEs are not
hydrocarbons, they are structurally similar to the paraffin hydrocarbons present in
diesel, because both paraffins and FAMEs contain straight chains of carbon
molecules. For this reason, M. methanica is an organism that has the capability
to potentially contribute to the production of biodiesel FAMEs.
Considering the factors of price and availability, the future of transportation
worldwide will soon be dependent on the production of renewable fuels.
Currently ethanol is being mixed with gasoline (up to 10 percent) and biodiesel is
being mixed with diesel fuel. While these practices are more environmentally
friendly than using pure fossil fuel and meet current government regulations, they
do not completely reduce dependence on foreign oil or provide a completely
renewable fuel source. However, Solazyme, a company that produces 100
percent algal derived biodieselCIT, has made a promising advancement.
Recently, Solazyme provided 80,000 liters of biodiesel to the United States Navy.
This transaction was the world’s largest delivery of non-ethanol biofuel that was
100 percent microbial derived. The Navy vessel USS Ford successfully voyaged
from Everett, WA, to San Diego, CA, using algal-derived biodiesel. The vessel
did not require engine or fueling dock modifications (Solazyme, Inc., 2012). The
fact that the US Navy has proven that biodiesel is an effective, renewable
alternative to fossil fuel derived diesel paves the way for widespread use of
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biodiesel in the United States and worldwide (Solazyme, Inc., 2012). Even more
recently, Solazyme has produced SoladieselBD®, a fatty acid methyl ester (FAME)
based fuel. This product has been sucecssfully tested for thousands of miles in
unmodified vehicles, and has also demonstrated better cold temperature
properties than any other commercially available biodiesel
(Solazyme, Inc., 2013).
1.3 Methane metabolism
As previously mentioned, the defining characteristic of methanotrophic bacteria is
the unique ability to utilize methane or methanol as a sole source of carbon and
energy. This metabolic capability is because of the enzyme methane
monooxygenase (MMO), which is the initial enzyme required for methane
oxidation. The enzyme oxidizes methane by splitting a dioxygen (O2) molecule
and incorporating one oxygen atom into methane to form methanol. The other
oxygen atom is reduced to form water. Two different variations of this enzyme
have been identified: a particulate or membrane bound methane
monooxygenase (pMMO), and a soluble methane monooxygenase (sMMO)
(Hanson and Hanson 1996, Trotsenko and Murrell 2008). The enzyme pMMO
is ubiquitous among methanotrophs, and has been purified from alpha and
gamma Proteobacteria. However, sMMO has only been detected in a few
methanotrophic bacteria, and occurs only in bacteria that also express pMMO.
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The enzyme pMMO is associated with the intracytoplasmic membranes (ICM)
that are characteristic of methanotrophs.
The membranes of methanotrophs also differ between Type I and Type II
organisms. The membranes of Type I methanotrophs are arranged in stacks of
vesicular disks extending horizontally across the cell. In contrast, membranes in
Type II methanotrophs are arranged along the cell wall. Regardless of
arrangement, the intracytoplasmic membrane and the associated pMMO provide
an extensive surface area for methane oxidation.
Growth conditions also influence the formation of extensive membranes.
Methane – utilizing bacteria exhibit enhanced formation of intracytoplasmic
membranes (ICMs), while microbes supplied with methanol did not possess
abundant ICM (Best and Higgins 1981). Additionally, methanotrophs that are
exposed to oxygen-limited conditions fail to produce extensive ICM
(Scott et al. 1981). A lack of ICM formation has also been observed in growth
conditions that are devoid of copper (Prior and Dalton 1985). This observation
has lead to the investigation of the role of copper in ICM formation as well as in
pMMO. Because pMMO is an integral membrane enzyme, efforts to purify and
characterize this enzyme have been impeded. However, the crystal structure of
pMMO has been determined, revealing that it is composed of three subunits:
pmoA, pmoB, and pmoC (Lieberman and Rosenzweig 2005; Hakemian et al.
2008). The di-copper center of pMMO has been proven to be the catalyst for the
enzyme (Balasubramanian et al. 2010).
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In addition to these recent findings, copper-binding molecules called
methanobactins have been discovered (Lidstrom 2006). Methanobactins
sequester copper from the environment and are similar to siderophores.
Methanobactins are thought contribute to the regulation of methane
monooxygenase expression, the activity of pMMO, copper uptake, and the
protection of methanotrophic organisms from copper toxicity
(Kim et al. 2004, Balasubramanian and Rosenzweig 2008). Copper
concentration also plays an important role in regulating the expression of each
enzyme in cells capable of expressing both pMMO and sMMO. When copper is
present in concentrations of approximately 4 µM, pMMO is expressed and
extensive intracytoplasmic membranes develop. Alternatively, when copper
concentrations are limited to less than 0.8 µM, sMMO becomes expressed
(Nielsen et al. 1997, Prior and Dalton 1985, Choi et al. 2003).
The enzyme sMMO has a very broad substrate range and has been
purified from both alpha and gamma proteobacteria (Lipscomb 1994). This
enzyme has the ability to oxidize or hydroxylate a variety of aromatic, branched,
halogenated, and aliphatic straight-chained hydrocarbons. For this reason,
methanotrophic organisms have become an important factor in the
bioremediation of many toxic hydrocarbons (Hanson and Hanson 1996). In
contrast to pMMO, sMMO has a hydroxo-bridged di-iron center and obtains
reducing power from NADH (Lipscom 1994). The enzyme sMMO has three main
components. The first is a hydroxylase, which is composed of a non-heme iron
14
center and three polypeptides. The second component is component B, which
has no cofactors. Finally, the third component is a reductase that contains a
Fe2S2 cluster and FAD (Lipscomb 1994).
After the initial oxidation of methane to methanol by MMO, methanol is
oxidized to formaldehyde by a quinoprotein methanol dehydrogenase (MDH).
MDH is composed of two subunits that are the cofactor pyrroloquinoline quinone
(PQQ), and the subunit mxaFI (Goodwin and Anthony 1998). MDH and the two
related cytochromes (CL and CH) are located in the periplasm of the cell. MDH
transfers electrons to cytochrome CL, which is a specific electron acceptor for this
particular dehydrogenase. Cytochrome CL is then oxidized by cytochrome CH,
and the oxidation of methanol to formaldehyde is completed
(Hanson and Hanson 1996).
Formaldehyde is the central intermediate of methane metabolism. At this
point in the metabolic pathway, formaldehyde is either oxidized to CO2 or
assimilated into cellular material. The enzyme formaldehyde dehydrogenase
(FaDH) converts formaldehyde to formate. This oxidation step is responsible for
producing the majority of reducing power required for methane metabolism. In
the dissimilatory metabolic step, the formate produced from the oxidation of
formaldehyde is oxidized to carbon dioxide by an NAD-dependent formate
dehydrogenase (Hanson, 1996). Alternatively, the assimilation of formaldehyde
employs the use of one of two separate pathways: the ribulose monophosphate
(RuMP) pathway or the serine pathway. The RuMP pathway, which was
15
described by Quayle and his colleagues, (Johnson and Quayle, 1965) is present
in Type I methanotrophs such as Methylomonas, methylomicrobium,
Methylobacter, Methylosphaera, and Methylocaldum. This pathway requires 3
moles of formaldehyde to form glyceraldehyde-3-phosphate, which is assimilated
into cellular material.
Alternatively, the serine pathway that is present in Type II methanotrophs
such as Methylosinus and Methylocystis, requires 1 mole of carbon dioxide and 2
moles of formaldehyde to form the three carbon intermediate serine. Serine then
undergoes several transformations, one of which is the conversion of 2-glycerate
to 2-phosphoglycerate, which is assimilated into cell material
(Hanson and Hanson 1996). To complete the cycle, malyl-CoA is cleaved to
form two, 2- carbon compounds: glyoxylate and acetyl-CoA. Glyoxylate is
converted to glycine and acetyl-CoA is transformed to regenerate the second
molecule of glyoxylate (Chistoserdova and Lidstrom 2013).
.
16
CHAPTER TWO
Materials and Methods
2.1 Materials
2.1.1 Microbial Cultures
Methylomonas methanica was used for all pure culture experiments. This
bacterium is a Type I methanotroph that was obtained from the American Type
Culture Collection (ATCC 51626).
2.1.2 Growth Medium
The growth medium used for this study was Nitrate Mineral Salts (NMS) medium
as described by ATCC. The components are (g/L): MgSO4 7H2O (1), KNO3 (1),
Na2HPO47H2O (0.717), KH2PO4 (2.72), CaCl32H20 (0.13), and a trace element
solution (1ml/L). Nanopure water (Barnstead Nanopure Diamond) was used for
medium preparation.
The components of the trace elements solution (TES) were (g/L):
FeSO47H2O (0.2), H3BO3 (0.03), CoCl26H2O (0.02), CuSO4 5H2O (0.03),
ZnSO47H2O (0.01), MnCl24H2O (0.003), Na2MoO42H2O (0.003), and
NiCl26H2O (0.002). After mixing, the trace elements mixture was sterilized using
a vacuum filtration unit equipped with Whatman No. 1 filter paper, 11µm pore
size. The NMS medium components were added to 910 mL of nanopure water to
prepare a 1x concentrated solution to ensure that the media components would
17
be in the desired concentration after inoculation. The medium was mixed for 20
minutes or until all components were dissolved. The pH of the medium was
adjusted to 6.8. Thirty mL of the growth medium were dispensed into 165 mL
glass serum bottles. The serum bottles were closed with a butyl rubber stopper,
sealed with an aluminum crimp cap and autoclaved for 20 min at 121°C. After
cooling, 0.1mL of filtered trace element solution was aseptically added to each
serum bottle with a sterile 1 mL syringe and needle prior to the start of the
experiment.
2.2 Methods 2.2.1 Growth of M. methanica
Growth curve, temperature, copper concentration, and enrichment experiments
were conducted in the 165 mL glass serum bottles containing growth medium as
described above. Using a sterile 5 mL syringe and a 21 gauge needle, each
serum bottle was aseptically inoculated with 3 mL of M. methanica grown to a
cell density of 0.4 (OD600nm). After inoculation, the headspace of each bottle was
aseptically adjusted to have an approximate methane gas composition of 15%
methane by removing 20 mL of the original headspace using a sterile 10 mL
syringe, and injecting 20 mL of methane gas (Airgas, commercial grade) which
was obtained from a pressurized 275 mL glass bottle. All serum bottles were
incubated in a New Brunswick Innova 40 shaking incubator at 210 rpm.
18
2.2.2 Headspace Analysis
Headspace analysis was carried out by initially measuring the headspace
pressure of the serum bottle using a manometer (Dwyer Series 477). A 0.5 mL
sample of the headspace gas was removed from the serum bottle before cell
density was measured using a 1 mL tuberculin syringe (BD, Franklin Lakes, NJ)
and injected into a gas chromatograph (Agilent 7890A) equipped with a HP-
PLOT Molesieve column (Agilent J&W, 30 m, 0.32 mm, 3 µm) operating as
follows: inlet at 250°C split 10:1, 8.6 psi, total flow 102 mL/min, set to a constant
pressure of 8.9 psi, with the oven temperature at 50°C, held for 1.5 minutes then
to 150°C at 50°C/min and held for 2 minutes. The thermal conductivity detector
(TCD) was held at 250°C with a reference flow 20 mL/min and makeup flow at 2
mL/min. Helium was used as the carrier gas at a flow rate of 30 mL/min.
2.2.3 Lipid Extraction and Conversion to Fatty Acid Methyl Esters (FAMES) Samples were prepared for lipid extraction by placing the liquid culture in a 50
mL plastic centrifuge tube (VWR) and centrifuging at 4500 x g for 20 minutes in
order to pellet cells. The pellet was then washed with 10 mL deionized water two
times to remove residual salts. The washed pellet was then placed in a methanol
washed glass test tube with a threaded cap. Phospholipid fatty acids obtained
were converted to FAMEs using the MIDI, INC esterification technique. This
conversion process used the following reagents: Reagent 1 consisted of 45 g
19
sodium hydroxide, 150 mL methanol, and 150 mL distilled water; Reagent 2
consisted of 325 mL 6 N hydrochloric acid, 275 mL methyl alcohol; Reagent 3
consisted of 200 mL hexane, 200 mL methyl tert-butyl ether; and, Reagent 4
consisted of 10.8 g sodium hydroxide in 900 mL Nanopure water. This
esterification technique was performed on each sample separately by placing the
cell pellet into a methanol washed glass test tube with a threaded cap. Then, the
first step, saponification, was initiated by placing 1mL of reagent 1 into the test
tube using a glass pipette. The tube was sealed, vortexed for 5-10 seconds,
placed in a 100°C water bath for 5 minutes, vortexed for 5-10 seconds, and
returned to the water bath for 25 minutes. After cooling, the next step,
methylation, began by adding 2 mL of Reagent 2 to the contents of the test tube,
vortexing for 5-10 seconds, and heating in an 80°C water bath for 10 minutes.
The tube was then cooled rapidly in ice. The third step, extraction, began with
the addition of 1.25 mL of Reagent 3. The test tube was then placed on a clinical
rotator for 10 minutes. Phase separation was observed at this point, and the
lower phase was removed with a glass pipette and discarded. The remaining
phase was washed with 3 mL of Reagent 4, and placed on a clinical rotator for 5
minutes. Two-thirds of the top phase was removed with a glass pipette, placed
into a GC vial, and sealed with a threaded cap.
20
2.2.4 Quantification of FAMES The FAMEs obtained from each experiment were analyzed using a gas
chromatograph (Agilent 7890A) equipped with a flame ionization detector (FID)
and a HP INNOWAX column (Agilent) 30 m x 0.25 mm x 0.25 µm). Operating
conditions were as follows: inlet temperature 260°C, pressure 27.239 psi, total
flow 47.1 mL/min, septum purge 3 mL/min, split ratio 20:1, initial temperature of
140°C held for 5 minutes, increasing in increments of 4°C per minute, up to 240
°C and held for 5 minutes. A 0.2 µL sample was injected onto the column by an
autosampler (Agilent 7683b). Underivitized lipid standards (GLC Reference
Standard 20A, Nu-Chek Prep Inc.) were converted to FAMEs using the MIDI
esterification technique. The lipids present in the standard mixture were as
follows (each lipid as a percentage of the total lipid weight): C14:0/2.0,
C16:0/30.0, C16:1/3.0, C18:0/14.0, C18:1/41.0, C18:2/7.0, C18:3/3.0. Lipids
from 13.2 mg of this underivitized lipid mixture were extracted, and the resulting
FAMEs were diluted in hexane. Dilutions of 1:1, 1:10, 1:100, and 1:1000 were
prepared in GC vials. A standard curve was then constructed from these
standards (Appendix A, Figure A1).
2.3 Experimental Design 2.3.1 Growth curves at varying temperatures
The lipid composition of M. methanica was evaluated at the three different
temperatures of 25°C, 30°C, and 35°C. In order to correlate the growth versus
21
lipid production, a growth curve of M. methanica was performed at each
temperature. Each growth curve consisted of twelve time points, and each point
had triplicate serum bottles, containing the previously described medium, for
sampling. Each serum bottle was inoculated with 3 mL of a culture of M.
methanica grown to a cell density of 0.4 at OD600nm. Upon inoculation, the
amount of methane in the headspace of each bottle was analyzed via TCD as
described. All bottles were placed in an incubator set to the appropriate
temperature and shaken at 210 rpm. At each time point, three serum bottles
were removed. The headspace of each bottle was sampled to determine final
methane consumption, and the absorbance of each sample was determined by
placing 2 mL of sample into a cuvette. The absorbance was then analyzed using
the OD600 setting on a biophotometer (Eppendorf).
2.3.2 Dry weight versus Absorbance The amount of dried biomass produced at a particular absorbance was
measured to relate absorbance to dry biomass in future experiments. The dried
biomass from growth curves at absorbances (OD600) of 0.1, 0.2, 0.3, 0.4, 0.5, 0.6,
0.7, and 0.8 were evaluated. An initial headspace analysis was performed to
measure the methane content. The serum bottles were then placed in an
incubator set to 30°C, 210 rpm. Three bottles were chosen for each time point.
At each sampling timepoint, a headspace sample was taken followed by an
absorbance reading (OD600). These sampling techniques have been previously
described above. Each bottle was then emptied into separate 50 mL centrifuge
22
tubes that had been dried in an oven set to 80°C. After drying 48 hours, the
empty tubes and caps were weighed on an analytical balance. Each sample was
placed in its respective pre-weighed tube and centrifuged at 4500 rpm x g for 20
minutes (Eppendorf). The supernatant was then removed, and the pellet was
washed twice with 10 mL of deionized water. The tube, cap, and pellet were
then dried in an 80°C oven for 48 hours. The weight of the dried biomass was
then measured using an analytical balance.
2.3.3 FAME profiles at various temperatures In order to compare the FAME profiles produced by M. methanica at the
temperatures of 25°C, 30°C, and 33°C, the organism was grown to the same
absorbance at each temperature. The FAME profile during log and stationary
phases of growth was also evaluated. Thus, two cell densities
(absorbance OD600nm) for each temperature were sampled in triplicate. These
points were 0.4 OD (late-exponential phase) and 0.7 OD (stationary phase) for
the three temperatures. The sampling time for both growth phases at each
temperature was determined based on the results of the temperature growth
curves. Triplicate serum bottles for each sampling point were inoculated with 3
mL of a culture of M. methanica grown to an absorbance of 0.4 at OD. The initial
methane content in the headspace of each bottle was recorded, and the bottles
were placed in an incubator set to 25°C, 30°C, or 33°C at 210 rpm. Three bottles
were then sampled at the appropriate growth phase for each respective
temperature. A final headspace analysis was used to measure methane
23
consumption prior to the absorbance reading. Each sample was then placed in a
50 mL centrifuge tube and centrifuged for 20 minutes at 4500 rpm x g. The
supernatant was removed and the remaining pellet was washed twice with 10 mL
of deionized water to remove residual salts. The cell pellet for each sample was
then transferred to a methanol-washed glass test tube. The lipids from each
sample were extracted and converted to FAMEs using the MIDI, INC
esterification technique described previously. The resulting FAMEs were placed
in a GC vial and quantified using a gas chromatograph equipped with a flame
ionization detector (FID).
3.1 Effect of copper on quantity of lipids produced by M. methanica After evaluating the lipid profiles of M. methanica at 25°C, 30°C, and 33°C, the
growth temperature of 25°C was selected because it resulted in the best ratio of
biodiesel-like lipids. The late-exponential growth phase was also found to
produce the greatest amount of these lipids based on biomass production and
methane consumption. M. methanica was then grown under these selected
conditions while being exposed to different concentrations of copper. The lipid
production at the concentrations of 0, 0.5, 5, 10, 20, and 50 µM CuSO4 was
evaluated. Triplicate serum bottles per concentration were inoculated using the
previously described method. An initial headspace sample was taken from each
serum bottle to measure the initial amount of methane present. All bottles were
then incubated at 25 °C for 20 hours. A final headspace analysis was performed
24
to determine the amount of methane consumed during growth. An absorbance
reading was then taken to ensure that the samples were in exponential phase.
Samples were then converted to FAMEs and analyzed as previously described.
3.2 Growth of M. methanica under enhanced conditions in 2-L flask
M. methanica was then grown at the optimized conditions of 25°C, late-
exponential phase, and 5 µM copper concentration by adding 445.5 mL of
medium, pH 6.8, to a 2 L flask. The flask was sealed with a butyl-rubber stopper
and aluminum crimp cap and autoclaved for 20 min at 121°C. After cooling, 2.25
mL of a 1 mM solution of CuSO4 was added to the medium after it was sterilized
using a 0.2-µm syringe filter and aseptically added to the 2 L flask. Then, 0.5 ml
of filter-sterilized trace element solution was also added aseptically with a
separate sterile 1 mL syringe and needle. The bottle was inoculated with 50 mL
of M. methanica at a cell density of 0.4 OD. Finally, 265 mL of headspace was
removed with a sterile 30 ml syringe and needle and replaced with 265 ml of
methane gas. The headspace of the bottle was analyzed for methane content
before incubation. The bottle was placed in a 25°C incubator and shaken at 210
rpm for 20 hours. A final headspace analysis was performed prior to the
evaluations of cell density, FAME profile and quantity of FAMEs produced using
the procedures described previously.
25
CHAPTER THREE
Results and Discussion
3.1 Research Objective
Methane gas is the second most abundant greenhouse gas in the atmosphere,
and is also considered the most inert hydrocarbon. The energy stored in the
four-carbon/hydrogen bonds of one molecule of methane has the potential to be
converted into a readily useable form of energy. Traditional catalysts that break
this bond (Balasubramanian et al. 2010) require high temperatures to be
effective. Alternatively, methanotrophic bacteria use the enzyme methane
monooxygenase to oxidize methane gas as a sole carbon and energy source,
which is assimilated into cell material, specifically extensive intracytoplasmic
membranes. These intracytoplasmic membranes are composed of PLFAs that
are similar to the composition of soy biodiesel.
The overall aim of this research was utilize the methane-oxidizing activity
of the methanotrophic bacterium Methylomonas methanica to convert methane
gas into biodiesel-like lipid components. The first objective for this research was
to evaluate the methane-oxidizing capabilities of M. methanica, in conjunction
with the potential for this organism to produce a lipid profile similar to soy
biodiesel. The second objective was to evaluate whether manipulating the
growth conditions of the bacterium could enhance the lipid profile of M.
methanica. The effect of the growth phase and growth temperature on the types
26
of lipids produced by this organism was evaluated. After evaluating the effect of
temperature, the effect of copper concentration on the amount of lipids produced
was evaluated. Finally, the third objective was to combine the enhanced
conditions of growth phase, temperature, and copper concentration to evaluate
the potential production of biodiesel-like lipids by M. methanica in a larger bench-
scale system.
3.2 Potential production of biodiesel lipids from methane gas
The potential for M. methanica to utilize methane gas as a sole carbon and
energy source and assimilate this methane into biodiesel-like lipids was
evaluated. Based on the research conducted by Guckert et al. (1991), M.
methanica produces a FAME profile that consists of individual FAMEs that are
14:0, 16:0, and 16:1 carbons in length, with the C16:1 FAME composing the
majority of the profile. After obtaining the culture (ATCC 51626), M. methanica
was grown at 30°C with a headspace consisting of 15% methane gas. Shaking
the culture during growth maintained an even distribution of methane gas in the
system. Cultures grown in the absence of shaking formed a thin pellicle on the
surface of the growth medium; however, these samples failed to produce a
measureable amount of biomass.
M. methanica cultures were grown for 48 hours, and the consumption of
methane was also determined (Figure 1). The data in this figure indicates that M.
methanica consumed 0.756 mmol of methane gas over 48 hours. The weight of
27
dried biomass was measured. PLFAs were also extracted from these samples
and converted to fatty acid methyl esters (FAMEs). The amount of total FAMEs
can be observed in Figure 1, which indicates that these FAMES are produced by
this culture of M. methanica. The data obtained demonstrates that M. methanica
is in fact capable of consuming methane gas and producing lipids similar to those
present in biodiesel.
28
Figure 1. The quantities of biomass and FAMES produced during growth of M. methanica at 30°C and the quantity of methane gas consumed during growth.
29
3.3 The effect of growth temperature on FAME ratios
3.3.A Determination of growth rates and methane consumption
Previous research has established that bacteria must maintain a fluid membrane
in order for cellular processes to function normally. At higher temperatures,
membrane lipids tend to be predominantly saturated. When growth temperatures
are lowered, the cell membrane undergoes a transition from the disordered state
of predominantly saturated membrane lipids to an ordered, nonfluid state of
predominantly unsaturated membrane lipids (de Mendoza and Cronan 1983). M.
methanica cultures were grown at 20°C, 25°C, 30°C, 33°C and 35°C to evaluate
the effect temperature had on growth rate, maximum cell densities, and the
FAME composition. Growth was not observed at 20°C, and minimal growth was
observed at 35°C. At 35°C, signs of clumps were observed. The growth profiles
at temperatures of 25°C, 30°C, and 33°C were chosen for evaluation.
In addition to evaluating the effect of temperature on membrane FAME
ratio, the effect of growth phase on the FAME ratio as well as the quantity of
FAMEs was investigated. Research conducted by Clark and Cronan (1996)
revealed that bacteria are capable of utilizing cellular lipids as a source of carbon
and energy through a process called β-oxidation (Clark and Cronan 1996). This
process is essentially the reverse of the assimilation of cellular lipids (Vance
2008). Based on this information, the hypothesis that bacterial cells will
metabolize cellular lipids when later stages of growth are reached can be formed.
Cells may also begin the process of β-oxidation when the original source of
30
carbon and energy, in this case methane, is depleted. Therefore, the FAME
profiles, as well as the quantity of FAMEs produced, were assessed at late-
exponential as well as stationary growth phases at each growth temperature.
In order to evaluate the conditions of temperature and growth phase, growth
curves were conducted for the temperatures of 25°C, 30°C, and 33°C. Methane
consumption during growth at each temperature was also monitored. Because
the growth phases of late-exponential and stationary were reached at different
times depending on the temperature, growth curves were deemed necessary. At
25°C, the doubling time of M. methanica was found to be 7.2 hours, and the
organism reached late-log phase at 20 hours and stationary phase at 38 hours
(Figure 2a). The amount of methane consumed at late log phase was 0.35
mmol, with 0.71 mmol being consumed at stationary phase (Figure 3a). At 30°C,
the doubling time of M. methanica was determined to be 6.5 hours, and the
organism reached late-log phase at 18 hours and stationary phase at 32 hours
(Figure 2b). The amount of methane consumed at late log phase was 0.44
mmol, with 0.68 mmol being consumed at stationary phase (Figure 3b). Finally,
the doubling time of M. methanica at 33°C was determined to be 4.2 hours, and
the organism reached late-log phase at 16 hours and stationary phase at 28
hours (Figure 2c). The amount of methane consumed at late log phase was 0.41
mmol, with 0.73 mmol being consumed at stationary phase (Figure 3c). These
data allowed samples for FAME analysis to be collected at the same cell density
and growth phase
31
Figure 2. Growth Profiles of M. methanica at (a) 25°C (b) 30°C and (c) 33°C
32
Figure 3. Methane consumption during growth at (a) 25°C (b) 30°C and (c) 33°C
33
3.3.B Normalization of cell density to weight of dried biomass
To evaluate the quantity of FAMEs produced in each sample, the weight of
overall biomass produced was required. Because each experiment was
performed in triplicate with each bottle being sacrificed per sample, the
determination of the dry weight of each sample was not feasible. Therefore, an
experiment to normalize cell density (OD600) to dry weight of biomass was
conducted.
M. methanica was grown at 30°C, and samples were collected at several
cell densities as measured by optical density. The weight of dried biomass (mg)
was measured for each cell density, and these data were used to create a
standard equation, shown in Figure 4, to calculate the weight of dried biomass
based on cell density.
34
Figure 4. Plot used to normalize dried biomass to cell density.
35
3.3.C The effect of temperature and growth phase on total FAME production
Cultures of M. methanica were grown to late-log and stationary phases at the
temperatures of 25°C, 30°C, and 33°C. The amount of methane consumed
during growth at each temperature was also monitored. The cell density of each
sample was measured, and the amount of total biomass produced was
calculated using the method described in section 3.3.B. PLFAs were extracted
from each sample for subsequent conversion to FAMEs. The FAME ratios as
well as the amount of each individual FAME produced were evaluated using pre-
determined FAME standards of known quantities. Figure 5 depicts the total
FAMEs extracted from each sample as a percentage of the calculated dried
biomass for each respective sample. The percentage of total FAMEs produced
was greater in late-exponential phase samples than stationary phase samples for
every temperature. The difference in the percentage of total FAMEs produced in
late-exponential phase at each temperature was not significant, however, the
amount of biomass produced at 33°C was significantly less than the amounts
produced at 25°C and 30°C.
36
Figure 5. The quantity of biomass and percentage total FAMEs produced by M.
methanica at late-exponential and stationary phases while growing at temperatures 25°C, 30°C, and 33°C.
37
3.3.D Effects of temperature and growth phase on the production of individual
FAMEs
In addition to the evaluation of total lipids in section 3.3.C, the quantity of the
individual FAMEs produced at each growth phase and temperature was
measured. The FAMEs produced by M. methanica are C14:0, C16:0, and C16:1.
The amount of these individual FAMEs produced at each growth phase and
temperature is depicted in Figure 6. Figure 6a indicates the amount of each
FAME (mg/L), as well as the amount of total FAMEs produced at late-exponential
and stationary phase at 25°C. The FAME C16:1 is the most abundant FAME in
this sample. Figure 6b represents the amount of each FAME (mg/L), as well as
the amount of total FAMEs produced at late-exponential and stationary phase at
30°C. Again, the FAME C16:1 is the most abundant FAME in this sample. Figure
6c provides the amount of each FAME (mg/L), as well as the amount of total
FAMEs produced at late-exponential and stationary phase at 33°C. Again, the
FAME C16:1 is the most abundant FAME in this sample.
To evaluate the FAME ratio for each growth phase and temperature, the
individual FAME were expressed as a percentage of the total FAMEs produced.
From the results in Figure 7, the temperature of 25°C in late-exponential phase
was found to produce the highest percentage of the C16:1 FAMEs. The lipid
yield based on the consumption of methane gas was also calculated (Table 2).
The assumption can be made that the majority of the carbon derived from
38
methane is assimilated into C16:1 FAMEs at the temperature of 25°C and late-
exponential growth phase.
39
Figure 6. The quantity of total and individual FAMEs produced by M. methanica while growing at
exponential and stationary phases, 25°C, 30°C, and 33°C
40
Figure 7. The ratios of FAME produced by M. methanica while growing at late exponential and
stationary phase and at the temperatures of 25°C, 30°C, and 33°C.
41
25°C 30°C 33°C
Late exponential 0.088756354 0.108959531 0.084710484
(+ or -) 0.00569291 0.012705172 0.005211611
Stationary 0.053562537 0.057619407 0.050389421
(+ or -) 0.006312137 0.006790114 0.001432639 Table 2. Lipid Yield of M. methanica at late-exponential and stationary phases, temperatures 25°C, 30°C, and 33°C (g FAMEs produced / g methane consumed)
42
3.4 The effect of copper concentration on quantity of lipids produced
After assessing the influence of growth temperature and growth phase on the
ratio of FAMEs, the effect of copper concentration on the quantity of FAMEs
produced was evaluated. Previous research with Methylococcus capsulatus
Bath indicated an increase in cell mass, abundance of intracytoplasmic
membranes, and pMMO when copper concentrations are 5-10 µM in the growth
medium (Collins et al. 1991). When cultures were grown without added copper
in the growth medium, minimal growth was observed. In addition, Brantner et al.
(1997) grow M. capsulatus Bath with copper concentration exceeding 1 µM, and
an increase in cell mass was observed. Because pMMO is an integral
membrane enzyme and the active site of this enzyme is a di-copper center, it
requires copper to function (Balasubramanian et al. 2010).
Based on this information, the hypothesis that growth medium copper
concentrations above 1 µM will influence the production of a greater quantity of
PLFAs because of the production of extensive intracytoplasmic membranes. The
growth of M. methanica at the copper concentrations of 0 µM, 1 µM, 5 µM, 10
µM, 20 µM and 50 µM was evaluated. No growth was observed at 50 µM, and
minimal growth and biomass production was observed at 20 µM. Growth
conditions for the evaluation of these copper concentrations were 25°C and late
exponential phase. These conditions were previously determined to influence
the desired ratio of FAMEs in experiment 3.3.
43
3.4.A Effect of copper on total FAME production
The effect of copper concentration on the total amount of FAMEs produced by M.
methanica at 25°C, late exponential phase was evaluated. Methane
consumption was also monitored for each sample. The culture was grown in
media that had concentrations of 0 µM, 1 µM, 5 µM, 10 µM, 20 µM and 50 µM.
The actual concentration of the 0 µM sample was determined to be 0.5 µM due
to pre-existing copper concentrations in the trace element solution that is
essential for the growth of this organism. This was taken into account when
determining the copper concentrations listed above. As previously mentioned,
no growth was observed at 50 µM, therefore PLFAs were not extracted from this
sample. Based on the growth curve of M. methanica at 25°C, each copper
concentration culture was grown for twenty hours, which coordinates with late-
exponential phase. The dry weight of each culture was measured based on cell
density (3.3B). PLFAs were extracted from each sample and converted to
FAMEs. The FAME amount (mg/L) is shown in Figure 8. The FAME yield based
on the amount of methane gas consumed was also calculated for each
concentration (Table 3). Although there is no significant difference in the amount
of FAMEs produced at each concentration, the greatest amount of biomass was
produced at the 5 µM concentration. The ratios of the amount of each individual
FAME in each sample were also evaluated. These ratios can be observed in
Figure 10. At the concentration of 5µM, the largest percentage of C16:1 FAMEs
44
were produced. The amount of C16:1 FAMES at this concentration was 11 mg/L
(Figure 9). Based on these observations, predictions that the copper
concentration of 5 µM influences the overall quantity of C16:1 FAMEs can be
made.
45
Figure 8. The amount of total biomass and percentage of total FAMEs produced when M. methanica was grown at different copper concentrations.
46
Figure 9. The quantity of total and individual FAMEs produced by M. methanica while growing at copper concentrations 0 µM, 1 µM, 5 µM, 10 µM, 20 µM
47
Figure 10. The ratios of FAMEs produced by M. methanica when grown with different copper concentrations at 25°C during late-exponential phase.
48
Table 3. Lipid yield of M. methanica at different copper concentrations. Culture conditions are 25°C, late-exponential phase
3.5 Growth of M. methanica under optimized conditions in a 2 bench-scale reactor. The potential for enhanced biodiesel lipid production on a larger scale was
evaluated by growing M. methanica in a 2 L flask under the optimized conditions
of 25°C, 5 µM copper concentration, and at late-exponential phase. In this
system, the volume of culture medium was 500 mL in comparison to the 33 ml
volume that was used for all other experiments in this study. The percentages of
methane and inoculum, 15% and 10% respectively, remained the same as used
previously. Figure 11 shows the amount of total FAMEs produced and the FAME
composition. The FAME C16:1 comprises 70% of the weight of total FAMEs
produced, with C14:0 and C16:0 comprising 21% and 9%, respectively. Table 4
has the lipid yields for this system and the carbon balance.
0µM 1µM 5µM 10µM 20µM
Lipid Yield (g FAME/ g methane)
0.063363414 0.05205846 0.057688649 0.078744002 0.05958257
(+ or -) 0.010811549 0.002326936 0.002019951 0.0016869 0.012178364
49
Figure 11. Amount of total lipids and percentage of individual lipids produced in 2 liter flask.
50
Lipid yield g lipids/g methane 10.20507243
g carbon (Methane consumed) g carbon i(n lipids)
carbon balance 0.006836412 0.003840073 Table 4. Lipid yield and carbon balance in 2 liter flask
Figure 12. Ratio of individual lipids in 2-L reactor
51
Figure 13. The quantities of biomass and FAMES produced during growth of M.
methanica at 25°C, late-exponential phase, 5 µM copper and the quantity of methane gas consumed during growth.
52
CHAPTER FOUR
Conclusion
The purpose of this research was to evaluate the potential of a methanotrophic
bacterium to grow on methane and produce fatty acids methyl esters (FAMES)
similar to those found in biodiesel fuel. If this bacterium could produce significant
quantities of FAMEs from methane then this could be a possible mechanism to
convert this from of natural gas to a liquid transportation fuel.
Methylomomas methanica was chosen as the methanotroph for this
research project because it is a Type I methanotroph that produces FAMEs
similar to those found in biodiesel. When grown on methane, M. methanica
produced more biodiesel-like FAMEs during late-exponential growth phase.
Although more biomass was observed during stationary growth phase, a
significantly smaller amount of FAMEs were produced. Therefore, exponential
phase was determined to be better suited for FAME production. Because the
production of C16:1 FAMEs was the focus of this research, the growth
temperature of 25°C was chosen for further study. This temperature produced
the highest ratio of C16:1 FAMEs (66.7%). The effect of copper on FAME
production was then evaluated at 25°C, late-exponential phase. The highest
ratio of C16:1 FAMEs was observed at the copper concentration of 5 µM.
Finally, M. methanica was grown in a 2-L flask in order to evaluate bacterial
growth on a larger scale. The growth conditions of 25°C, late-exponential phase,
and 5 µM copper concentration yielded 186 mg/L total biomass, 9 mg/L total
53
FAMEs, and 6.3 mg/L C16:1 FAMEs. C16:1 FAMEs comprised 70% of the total
FAMEs produced during growth in the 2-L flask. Based on these results, M.
methanica was determined to be a potential producer of C16:1 biodiesel-like
FAMEs, and this organism has the potential to convert methane gas into
biodiesel-like products.
54
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APPENDICES
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Appendix A
Standards for Methane and FAME Measurement
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